Co-localization of mitochondria with chloroplasts is a light-dependent reversible response

Co-localization of mitochondria with chloroplasts in plant cells has long been noticed as beneficial interactions of the organelles to active photosynthesis. Recently, we have found that mitochondria in mesophyll cells of Arabidopsis thaliana expressing mitochondrion-targeted green fluorescent protein (GFP) change their distribution in a light-dependent manner. Mitochondria occupy the periclinal and anticlinal regions of palisade cells under weak and strong blue light, respectively. Redistributed mitochondria seem to be rendered static through co-localization with chloroplasts. Here we further demonstrated that distribution patterns of mitochondria, together with chloroplasts, returned back to those of dark-adapted state during dark incubation after blue-light illumination. Reversible association of the two organelles may underlie flexible adaptation of plants to environmental fluctuations.Key words: Arabidopsis thaliana, blue light, chloroplast, green fluorescent protein, mesophyll cell, mitochondrion, organelle positioningHighly dynamic cell organelles, mitochondria, are responsible not only for energy production, but also for cellular metabolism, cell growth and survival as well as gene regulations.^1,2 Appropriate intracellular positioning and distribution of mitochondria contribute to proper organelle functions and are essential for cell signaling.^3,4 In plant cells operating photosynthesis, the co-localization of mitochondria with chloroplasts has been a well known phenomenon for a long period of time.^5,6,7 Physical contact of mitochondria with chloroplasts may provide a means to transfer genetic information from the organelle genome,⁸ as well as to exchange metabolite components; a process required for the maintenance of efficient photosynthesis.^9,10,11Using Arabidopsis thaliana stably expressing mitochondrion-targeted GFP,¹² we have recently examined a different aspect of mitochondria positioning. Although mitochondria in leaf mesophyll cells are highly motile under dark condition, mitochondria change their intracellular positions in response to light illumination.¹³ The pattern of light-dependent positioning of mitochondria seems to be essentially identical to that of chloroplasts.¹⁴ Mitochondria occupy the periclinal regions under weak blue light (wBL; 470 nm, 4 µmol m⁻²s⁻¹) and the anticlinal regions under strong blue light (sBL; 100 µmol m⁻²s⁻¹), respectively. A gradual increase in the number of static mitochondria located in the vicinity of chloroplasts in the periclinal regions with time period of wBL illumination clearly demonstrates that the co-localization of these two organelles is a light-induced phenomenon.¹³In the present study, to ask whether the light-dependent positioning of mitochondria is reversible or not, a time course of mitochondria redistribution was examined transferring the sample leaves from light to dark conditions. The representative results (Fig. 1) clearly show that mitochondria re-changed their positions within several hours of dark treatment. Immediately after dark adaptation, mitochondria in the palisade mesophyll cells were distributed randomly throughout the cytoplasm (Fig. 1A and ^{ref. 13}). Chloroplasts were distributed along the inner periclinal walls and the lower half of the anticlinal walls. On the contrary, mitochondria accumulated along the outer (Fig. 1B) and inner periclinal walls when illuminated with wBL. Chloroplast position was also along the outer and inner periclinal walls. Many of the mitochondria located near the chloroplasts lost their motility. When wBL-illuminated leaves were transferred back to dark condition, the numbers of mitochondria and chloroplasts present on the periclinal regions began to decrease within several hours (Fig. 1C). After 10 h dark treatment, distribution patterns of mitochondria as well as chloroplasts almost recovered to those of dark-adapted cells (Fig. 1D).Open in a separate windowFigure 1Distribution of mitochondria and chloroplasts on the outer periclinal regions of palisade mesophyll cells of A. thaliana under different light conditions. Mitochondria (green; GFP) and chloroplasts (red; chlorophyll autofluorescence) were visualized with confocal microscopy after dark adaptation (A), immediately after wBL (470 nm, 4 µmol m⁻²s⁻¹) illumination for 4 h (B), after dark treatment for 6 h (C) and 10 h (D) following the 4-h wBL illumination, respectively. Bar = 50 µm.To our knowledge, this may be the first report that directly demonstrates that wBL regulates mitochondria and chloroplast positioning in a reversible manner, though the nuclei in A. thaliana leaf cells were also found to reverse their positions when transferred from sBL to dark conditions.¹⁵ Reversible regulation of organelle positioning in leaf cells should play critical roles in adaptation of plants to highly fluctuating light conditions in the nature. Since distribution patterns of mitochondria under wBL and sBL are identical to those of chloroplasts, we can assume that phototropins, the BL receptors for chloroplast photo-relocation movement,¹⁶ may have some role in the redistribution of mitochondria. On the other hand, we also found that red light exhibited a significant effect on mitochondria positioning (Islam et al. 2009), suggesting an involvement of photosynthesis. These possibilities are now under investigation.     

Chloroplasts autophagy during senescence of individually darkened leaves

We recently reported that autophagy plays a role in chloroplasts degradation in individually-darkened senescing leaves. Chloroplasts contain approximately 80% of total leaf nitrogen, mainly as photosynthetic proteins, predominantly ribulose 1, 5-bisphosphate carboxylase/oxygenase (Rubisco). During leaf senescence, chloroplast proteins are degraded as a major source of nitrogen for new growth. Concomitantly, while decreasing in size, chloroplasts undergo transformation to non-photosynthetic gerontoplasts. Likewise, over time the population of chloroplasts (gerontoplasts) in mesophyll cells also decreases. While bulk degradation of the cytosol and organelles is mediated by autophagy, the role of chloroplast degradation is still unclear. In our latest study, we darkened individual leaves to observe chloroplast autophagy during accelerated senescence. At the end of the treatment period chloroplasts were much smaller in wild-type than in the autophagy defective mutant, atg4a4b-1, with the number of chloroplasts decreasing only in wild-type. Visualizing the chloroplast fractions accumulated in the vacuole, we concluded that chloroplasts were degraded by two different pathways, one was partial degradation by small vesicles containing only stromal-component (Rubisco containing bodies; RCBs) and the other was whole chloroplast degradation. Together, these pathways may explain the morphological attenuation of chloroplasts during leaf senescence and describe the fate of chloroplasts.Key words: Arabidopsis, autophagy, chloroplast, dark treatment, leaf senescence, nutrients recyclingThe most abundant chloroplast protein is Rubisco, comprising approximately 50% of the soluble protein.¹ The amount of Rubisco decreases rapidly in the early phase of leaf senescence, and more slowly in the later phase. During senescence, chloroplasts gradually shrink and their numbers gradually decrease in mesophyll cells.^2,3 During leaf senescence, leaves lose approximately 75% of their Rubisco, while chloroplast numbers decrease by only about 15%.⁴ Previous studies showed chloroplasts localized within the central vacuole by electron microscopy, indicating chloroplast degradation in the highly hydrolytic vacuole.⁵ However, there was no direct evidence showing translocation of chloroplasts from the cytosol to the vacuole, and the mechanism of transportation was also unclear.Recent reverse genetic approaches are helping to elucidate the autophagy system in plants, which has a similar molecular mechanism as in yeast.^6–11 In Arabidopsis (Arabidopsis thaliana), atg mutants have phenotypically accelerated leaf senescence, insufficient root elongation in nutrient starvation condition and reduced seeds yields, therefore, autophagy is considered to be important for nutrient recycling especially nutrient starvation and senescence in plants.¹²In Arabidopsis, individually darkened rosette leaves (IDLs) exhibit enhanced senescence.¹³ Appling IDLs treatment as an experimental model of leaf senescence, we recently demonstrated that chloroplasts are degraded in two different pathways by autophagy, one for RCBs,^14,15 and one for whole chloroplast.¹⁶ Darkened leaves became pale in 3 to 5 days treatment, while illuminated parts normally grow in both wild-type and autophagy defective mutant, atg4a4b-1. Furthermore, genes specifically expressed during senescence, SAG12 and SEN1, were rapidly upregulated, meanwhile, photosynthetic genes, such as RBCS2B and CAB2B, were gradually downregulated. All analyzed ATG genes were also upregulated under IDL treatment, which suggests that autophagy is important in IDL senescence. It has been reported that approximately three quarter genes of upregulated in IDL were also upregulated in naturally senescing leaves, including the ATG genes.¹⁷ This suggests that the autophagy pathways used in IDLs are also used in naturally senescing leaves.Over the 5 day treatment period, chloroplasts of wild-type IDL shrink to approximately one third their original size. In atg4a4b-1, by contrast, chloroplasts shrinkage occurred immediately after the start of IDL treatment after which no further shrinkage was noted. While the shrunk chloroplasts in fixed cells of wild-type were still smooth and round, while wrinkly chloroplasts were observed in atg4a4b-1. At same time, in the living mesophyll cells of wild-type IDL, RCBs accumulated in the vacuole (Fig 1B). The shrinkage of chloroplasts may be due to the consumption of the chloroplast envelope by RCB formation. Immunological quantification of inner and outer envelope proteins might confirm this hypothesis. The chloroplast number was also gradually decreased in IDL of wild-type plants, but no decline in chloroplast number was noted in atg4a4b-1. Chloroplasts exhibiting chlorophyll auto-fluorescence were found in the vacuole of wild-type IDLs, but not in atg4a4b-1 IDLs. These results show that whole chloroplast degradation is also performed by autophagy. However, the transport pathway of whole chloroplasts into the vacuole remains unclear. The chloroplast, even in its shrunken state, is a large organelle, and the autophagosome, the carrier bodies of autophagy, which usually target small spherical organelles like mitochondria and peroxisomes, may be incapable of isolating large organelles. In the yeast autophagy system, specific cellular organelles and fractions are also transported via vacuolar membrane invagination using the microautophagy system.¹⁸ RCB uptake into the vacuole is termed macroautophagy, while larger organelles, such as chloroplasts, are engulfed in a process known as microautophagy. Whether there exists a molecular difference between these processes, or whether this is an arbitrary division based solely on the size of the consumed body is unclear.Open in a separate windowFigure 1Visualization of stroma-targeted DsRed and chlorophyll autofluorescence in living mesophyll cells of wild-type plants by laser-scanning confocal microscopy. A excised control leaf (A, Light) and an individually darkened leaf (B, IDL) from plants grown under 14 h-photoperiod condition and a leaf from whole-plant darkened condition (WD, C) for 5days were incubated with 1 µM concanamycin A in 10 mM MES-NaOH (pH 5.5) at 23C° for 20 h in darkness. Stroma-targeted DsRed appears green and chlorophyll fluorescence appears red. In merged images, overlap of DsRed and chlorophyll fluorescence appears yellow. Small vesicles with stromal-targeted DsRed, i.e. RCBs, can be found in the vacuole (A, B). In IDL (B), massive accumulation of stroma-targeted DsRed is entirely seen in the vacuolar lumen and chloroplasts losing DsRed fluorescence are found in some cells. Bars = 50 µm.Whole darkened plants exhibit retarded leaf aging, in contrast to the accelerated senescence in IDLs.¹³ Whole darkened plants suppress leaf senescence with the leaves retaining green color. After 5 days, in the mesophyll cells of whole darkened plants, any translocation of chloroplast components, stroma-targeted DsRed, RCBs, and whole chloroplasts, into the vacuole could hardly be detected (Fig. 1C). This suggests that autophagy is not induced by darkness alone, and is associated closely with senescence. ATG genes were downregulated in the whole darkened wild-type plants less than control plants during the treatment. Previous studies have shown that following about 5 day period of whole plant darkening, atg mutants lose their ability to protect themselves against photo-damage.⁷ Upon return to the light, these plant quickly undergo terminal photo-bleaching.Concentrations of chlorophyll, soluble protein, leaf nitrogen and Rubisco rapidly declined under IDL condition of both wild-type and atg4a4b-1. Considering the accumulated fluorescence of stroma-targeted Ds-Red in the vacuole and autophagy dependent size shrinkage of chloroplasts in IDL, in wild-type plants RCB autophagy appear to be responsible for a sizable proportion of chloroplast protein degradation. In atg4a4b-1 which cannot form RCBs, alternative degradation pathways must be upregulated, with chloroplast proteases the most likely candidates. Intriguingly, the decrease in Rubisco concentration proceeds at the almost identical rates in both wild-type and atg4a4b-1 plants, despite the different degradation pathways. It seems likely that the rate of Rubisco degradation may be regulated at an early step in the degradation pathway, by some, as yet unknown, factors.Chloroplasts appear to have the ability to control their volume during cell division, dividing and increasing their density up to the certain level,¹⁹ and transferring their cellular components between them via stromules.²⁰ How chloroplasts are able to regulate their volume remains unclear, but it seems likely that chloroplasts grow and divide, like any other bacteria, as long as sufficient resources remain in the environment, in this case the cell. Total chloroplast volume, therefore, may be limited by the availability of carbon, nitrogen, or other nutrients in the cell during leaf emergence. Chloroplasts may be also able to reduce and control their volumes during leaf senescence via multiple degradation pathways. Our next goal is to estimate the contribution of both RCBs and whole chloroplasts autophagy in chloroplast protein degradation during natural leaf senescence. Further investigations are required for understanding the specific molecular mechanisms of RCB production and whole chloroplast degradation.     

Endomembrane Ca2+-ATPases play a significant role in virus-induced adaptation to oxidative stress

Although the role of Ca²⁺ influx channels in oxidative stress signaling and cross-tolerance in plants is well established, little is known about the role of active Ca²⁺ efflux systems in this process. In our recent paper,¹⁷ we reported Potato Virus X (PVX)-induced acquired resistance to oxidative stress in Nicotiana benthamiana and showed the critical role of plasma membrane Ca²⁺/H⁺ exchangers in this process. The current study continues this research. Using biochemical and electrophysiological approaches, we reveal that both endomembrane P_2A and P_2B Ca²⁺-ATPases play significant roles in adaptive responses to oxidative stress by removing excessive Ca²⁺ from the cytosol, and that their functional expression is significantly altered in PVX-inoculated plants. These findings highlight the crucial role of Ca²⁺ efflux systems in acquired tolerance to oxidative stress and open up prospects for practical applications in agriculture, after in-depth comprehension of the fundamental mechanisms involved in common responses to environmental factors at the genomic, cellular and organismal levels.Key words: cytosolic calcium, reactive oxygen species, cross-tolerance, calcium pumpThe phenomenon of cross-tolerance to a variety of biotic and abiotic stresses is well-known.^1,2 Some of the demonstrated examples include the correlation between oxidative stress tolerance and pathogen resistance.^3–5 At the mechanistic level, changes in cytosolic Ca²⁺ levels [Ca²⁺]_cyt, have long been implicated as a quintessential component of this process.⁶ The rise in [Ca²⁺]_cyt is proven to be essential for the development of the oxidative burst required for triggering the activation of several plant defense reactions.^7,8 The observed elevation in H₂O₂ level is believed to result from Ca²⁺-dependent activation of the NADPH oxidase,⁸ which then causes a further increase in [Ca²⁺]_cyt via a positive feedback mechanism. This process is further accomplished by defense gene activation, phytoalexin synthesis and eventual cell death.⁹ Downstream from the stimulus-induced [Ca²⁺]_cyt elevation, cells possess an array of proteins that can respond to a message. Such proteins include calmodulin (CaM),¹⁰ Ca²⁺-dependent protein kinases¹¹ and CaM binding proteins.¹² Of note is that when Ca²⁺ channels are blocked, biosynthesis of ROS is prevented.¹³While the role of Ca²⁺ influx channels in oxidative stress signaling and cross-tolerance in plants is well established, little is known about the involvement of active Ca²⁺ efflux systems in this process. In contrast, in animal systems the essential role of re-establishing [Ca²⁺]_cyt to resting levels is widely reported. A sustained increase in [Ca²⁺]_cyt in the alveolar macrophage is thought to be the consequence of membrane Ca²⁺-ATPase dysfunction.¹⁴ In endothelial cells, inhibition of the Ca²⁺/Na⁺ electroneutral exchanger of the mitochondria was named as one of the reasons for [Ca²⁺]_cyt increases.¹⁵ A significant loss of the plasma membrane Ca²⁺-ATPase (PMCA) activity was reported in brain synapses in response to oxidative stress,¹⁶ suggesting that PMCA may be a downstream target of oxidative stress.In our recently published paper¹⁷ we reported the phenomenon of Potato Virus X (PVX)-induced acquired resistance to oxidative stress in Nicotiana benthamiana plants and showed the critical role of plasma membrane Ca²⁺/H⁺ exchangers in this process. Nonetheless, questions remain, is this transporter the only active Ca²⁺ efflux system involved in this process?In addition to Ca²⁺/H⁺ exchangers, active Ca²⁺ extrusion could also be achieved by Ca²⁺-ATPases. Two major types of Ca²⁺-ATPases that differ substantially in their pharmacology and sensitivity to CaM are known.¹⁸ Type P_2A pumps (also called ER-type or ECA^19,20) are predominantly ER-localized,¹⁹ although they are also present at other endomembranes (e.g., tonoplast and Golgi). Four members of this group have been identified in the Arabidopsis genome (named AtECAs 1 to 4).^18,21 These pumps lack an N-terminal autoregulatory domain, are insensitive to CaM and suppressed by cyclopropiazonic acid (CPA).¹⁹ P_2B (or ACA) pumps contain an autoinhibitory N-terminal domain that possesses a binding site for Ca²⁺-CaM.¹⁸ Ten members are known in Arabidopsis (termed AtACA1, 2, 4 and 7 to 13).²¹ Plant P_2B pumps are located at the plasma membrane²⁰ as well as in inner membranes such as tonoplast (e.g., ACA4), ER (e.g., ACA2) and plastids.^18,19 These pumps probably constitute the basis for precise cytosolic Ca²⁺ regulation; as the Ca²⁺ concentration increases, CaM is activated and binds to the autoinhibitory domain of the Ca²⁺ pump. This results in the activation of the pump.In our recent study,¹⁷ we found no significant difference between the purified plasma membranes fractions isolated from control and UV-treated tobacco plants (with or without PVX inoculation) either in the Ca²⁺-ATPase activity or in the Ca²⁺-ATPase expression level and its ability to bind CaM. This suggests that the plasma membrane P_2B type pumps (the only pump type known to be expressed at the plasma membrane) play no major role in removing excess Ca²⁺ from the cytosol under oxidative stress conditions. This led to an obvious question: what about endomembrane Ca²⁺-ATPases?To address this issue, microsomal membrane fractions were isolated from tobacco leaves in a manner previously described for plasma membrane fractions¹⁷ (Fig. 1A). Western blot and CaM overlay assays were then made to investigate the role of endomembrane P_2B Ca²⁺-ATPases in our reported phenomena of acquired resistance. The results show that the expression of the P_2B Ca²⁺ pumps in PVX-inoculated plants is significantly higher than in control plants (Fig. 1B), correlating well with the CaM overlay assay (Fig. 1C). As no difference was observed for the P_2B Ca²⁺-ATPase expression levels in the plasma membranes,¹⁷ the observed difference in the microsomal fractions of PVX-infected plants must be due to an increased expression of endomembrane P_2B Ca²⁺-ATPases. Given the fact that Ca²⁺ pumps have a high affinity for calcium, the observed increase in endomembrane P_2B-type Ca²⁺-ATPases expression in PVX-inoculated plants may be advantageous for more efficient Ca²⁺ removal from the cytosol into internal organelles.Open in a separate windowFigure 1Expression of P_2B Ca²⁺ in purified microsomal fractions from tobacco leaves. Measurements were undertaken C = mock controls; C-UV = mock controls treated with UV-light; PVX = PVX infected plants; PVX-UV = PVX inoculated plants treated with UV-light. (A) Coomassie Brilliant Blue-stained gel; (B) Protein blot immunostained with a non isoform-specific polyclonal antibody for P_2B Ca²⁺-ATPases; (C) CaM overlay assay.To decipher the possible role of P_2A Ca²⁺-ATPases in acquired resistance, a series of electrophysiological experiments were conducted using inhibitors of P_2A-type Ca²⁺-ATPases, such as thapsigargin (TG)²² and cyclopiazonic acid (CPA).²³ Ion-selective Ca²⁺ microelectrodes were prepared as described elsewhere in reference ²⁴ and ²⁵, and net Ca²⁺ fluxes were measured from tobacco mesophyll tissue following previously described protocols.¹⁷ Leaf pre-treatment for 2 h in either of these inhibitors dramatically suppressed the net Ca²⁺ efflux measured from tobacco mesophyll cells 2 h after UV light exposure (Fig. 2). Given the specificity of TG and CPA inhibitors for P_2A-type Ca²⁺-ATPases, these results strongly support a hypothesis that both endomembrane P_2A and P_2B Ca²⁺-ATPases play significant roles in plant adaptive responses to oxidative stress. This is achieved by removing excess Ca²⁺ from the cytosol.Open in a separate windowFigure 2Effect of known Ca²⁺-ATPase blockers on light-induced Ca²⁺ flux kinetics after 20 min of UV-C treatment. Leaf mesophyll segments were pre-treated in either 5 µM TG (thapsigargin) or 50 µM CPA (cyclopiazonic acid) for 1–1.5 h prior to exposure to UV-C light. Net Ca²⁺ fluxes were measured 2 h after the end of UV treatment. These were compared with two controls: (1) no pre-treatment/no UV exposure (closed circles) and (2) no pre-treatment/20 min UV exposure (open squares). Mean ± SE (n = 4 to 7).Combining these results with our previously reported observations in reference ¹⁷, the following model is proposed (Fig. 3). Oxidative stress (such as UV) causes increased ROS production in leaf chloroplasts, leading to the elevated [Ca²⁺]_cyt. Several Ca²⁺ efflux systems are involved in restoring basal cytosolic Ca²⁺ levels. Two of these, the plasma membrane Ca²⁺/H⁺ exchanger¹⁷ and endomembrane P_2A and P_2B Ca²⁺-ATPases (as reported in this study) are upregulated in PVX inoculated plants and contribute to the improved tolerance to oxidative stress. Overall, these findings highlight the potential role of Ca²⁺ efflux systems in virus-induced tolerance to oxidative stress in plants. This is consistent with our previous reports on the important role of Ca²⁺ efflux systems in biotic stress tolerance²⁶ and brings forth possibilities for genetic engineering of more tolerant plants by targeting expression and regulation of active Ca²⁺ efflux systems at either the plasma or endomembranes.Open in a separate windowFigure 3The proposed model of oxidative stress signaling and the role of Ca²⁺-efflux systems in acquired resistance and plant adaptation to oxidative stress.Overall, a better adaptation of virus-infected plants to a short wave UV irradiation as compared to uninfected controls may suggest that infection triggers common defense mechanisms that could be efficient against secondary unrelated stresses. This observation may lead to the development of novel strategies to protect plants against complex environmental stress conditions.     

Reactive oxygen species as transducers of sphinganine-mediated cell death pathway

Long chain bases or sphingoid bases are building blocks of complex sphingolipids that display a signaling role in programmed cell death in plants. So far, the type of programmed cell death in which these signaling lipids have been demonstrated to participate is the cell death that occurs in plant immunity, known as the hypersensitive response. The few links that have been described in this pathway are: MPK6 activation, increased calcium concentrations and reactive oxygen species (ROS) generation. The latter constitute one of the more elusive loops because of the chemical nature of ROS, the multiple possible cell sites where they can be formed and the ways in which they influence cell structure and function.Key words: hydrogen peroxide, long chain bases, programmed cell death, reactive oxygen species, sphinganine, sphingoid bases, superoxideA new transduction pathway that leads to programmed cell death (PCD) in plants has started to be unveiled.^1,2 Sphingoid bases or long chain bases (LCBs) are the distinctive elements in this PCD route that naturally operates in the entrance site of a pathogen as a way to contend its spread in the plant tissues.^2,3 This defense strategy has been known as the hypersensitive response (HR).^4,5As a lately discovered PCD signaling circuit, three connected transducers have been clearly identified in Arabidopsis: the LCB sphinganine (also named dihydrosphingosine or d18:0); MPK6, a mitogen activated kinase and superoxide and hydrogen peroxide as reactive oxygen species (ROS).^1,2 In addition, calcium transients have been recently allocated downstream of exogenously added sphinganine in tobacco cells.⁶Contrary to the signaling lipids derived from complex glycerolipid degradation, sphinganine, a metabolic precursor of complex sphingolipids, is raised by de novo synthesis in the endoplasmic reticulum to mediate PCD.^1,2 Our recent work demonstrated that only MPK6 and not MPK3 (commonly functionally redundant kinases) acts in this pathway and is positioned downstream of sphinganine elevation.² Although ROS have been identified downstream of LCBs in the route towards PCD,¹ the molecular system responsible for this ROS generation, their cellular site of formation and their precise role in the pathway have not been unequivocally identified. ROS are produced in practically all cell compartments as a result of energy transfer reactions, leaks from the electron transport chains, and oxidase and peroxidase catalysis.⁷Similar to what is observed in pathogen defense,³ increases in endogenous LCBs may be elicited by addition of fumonisin B1 (FB1) as well; FB1 is a mycotoxin that inhibits ceramide synthase. This inhibition results in an accumulation of its substrate, sphinganine and its modified forms, leading to the activation of PCD.^1,2,8 The application of FB1 is a commonly used approach for the study of PCD elicitation in Arabidopsis.^1,2,9–11An early production of ROS has been linked to an increase of LCBs. For example, an H₂O₂ burst is found in tobacco cells after 2–20 min of sphinganine supplementation,¹² and superoxide radical augmented in the medium 60 min after FB1 or sphinganine addition to Arabidopsis protoplasts (Fig. 1A). In consonance with this timing, both superoxide and H₂O₂ were detected in Arabidopsis leaves after 3–6 h exposure to FB1 or LCBs.¹ However, the source of ROS generation associated with sphinganine elevation seems to not be the same in both species: in tobacco cells, ROS formation is apparently dependent on a NADPH oxidase activity, a ROS source consistently implicated in the HR,^13,14 while in Arabidopsis, superoxide formation was unaffected by diphenyliodonium (DPI), a NADPH oxidase inhibitor (Fig. 1A). It is possible that the latter oxidative burst is due to an apoplastic peroxidase,¹⁵ or to intracellular ROS that diffuse outwards.^16,17 These results also suggest that both tobacco and Arabidopsis cells could produce ROS from different sources.Open in a separate windowFigure 1ROS are produced at early and long times in the FB1-induced PCD in Arabidopsis thaliana (Col-0). (A) Superoxide formation by Arabidopsis protoplasts is NADPH oxidase-independent and occurs 60 min after FB1 or sphinganine (d18:0) exposure. Protoplasts were obtained from a cell culture treated with cell wall lytic enzymes. Protoplasts were incubated with 10 µM FB1 or 10 µM sphinganine for 1 h. Then, cells were vacuum-filtered and the filtrate was used to determine XTT [2,3-bis-(2-methoxy-4-nitro-5-sulfophenyl)-2H-tetrazolium-5-carboxanilide, disodium salt] reduction as described in references ²⁸ and ²⁹. DPI was used at 50 µM. (B) H₂O₂ formation in Arabidopsis wt and lcb2a-1 mutant in the presence and absence of FB1. Arabidopsis seedlings were exposed to 10 µM FB1 and after 48 h seedlings were treated with DA B (3,3-diaminobencidine) to detect H₂O₂ according to Thordal-Christensen et al.³⁰It has been suggested that the H₂O₂ burst associated with the sphinganine signaling pathway leads to the expression of defense-related genes but not to the PCD itself in tobacco cells.¹² It is possible that ROS are involved in the same way in Arabidopsis, since defense gene expression is also induced by FB1 in Arabidopsis.⁹ In this case, it will be important to define how the early ROS that are DPI-insensitive could contribute to the PCD manifestation mediated by sphinganine.The generation of ROS (4–60 min) found in Arabidopsis was associated to three conditions: the addition of sphinganine (Fig. 1A), FB1 (Fig. 1A) or pathogen elicitors.¹⁵ This is consistent with the MPK6 activation time, which is downstream of sphinganine elevation and occurs as early as 15 min of FB1 or sphinganine exposure.² All of them are events that appear as initial steps in the relay pathway that produces PCD.In order to explore a possible participation of ROS at more advanced times of PCD progression, we detected in situ H₂O₂ formation in Arabidopsis seedlings previously exposed to FB1 for 48 h. As shown in Figure 1B, formation of the brown-reddish precipitate corresponding to the reaction of H₂O₂ with 3,3′-diaminobenzidine (DAB) was only visible in the FB1-exposed wild type plants, as compared to the non-treated plants. However, when lcb2a-1 mutant seedlings were used, FB1 exposure had a subtle effect in ROS formation. This mutant has a T-DNA insertion in the gene encoding subunit LCB2a from serine palmitoyltransferase (SPT), which catalyzes the first step in sphingolipid synthesis¹⁸ and the mutant has a FB1-resistant phenotype.² These results indicate that mutations in the LCB1¹ and LCB2a² genes (coding for the subunits of the heterodimeric SPT) that lead to a non-PCD phenotype upon the FB1 treatment, are unable to produce H₂O₂. In addition, they suggest that high levels of hydrogen peroxide are produced at advanced times in the PCD mediated by LCBs in Arabidopsis.Exposure of Arabidopsis to an avirulent strain of Pseudomonas syringae produces an endogenous elevation of LCBs as a way to implement defense responses that include HR-PCD.³ In this condition, we clearly detected H₂O₂ formation inside chloroplasts (Fig. 2A). When ultrastructure of the seedlings tissues exposed to FB1 for 72 h was analyzed, integrity of the chloroplast membrane system was severely affected in Arabidopsis wild-type seedlings exposed to FB1.² Therefore, we suggest that ROS generation-LCB induced in the chloroplast could be responsible of the observed membrane alteration, as noted by Liu et al. who found impairment in chloroplast function as a result of H₂O₂ formation in this organelle from tobacco plants. Interestingly, these plants overexpressed a MAP kinase kinase that activated the kinase SIPK, which is the ortholog of the MPK6 from Arabidopsis, a transducer in the PCD instrumented by LCBs.²Open in a separate windowFigure 2Conditions of LCBs elevation produce H₂O₂ formation in the chloroplast and perturbation in the membrane morphology of mitochondria. (A) Exposure of Arabidopsis leaves to the avirulent strain Pseudomonas syringae pv. tomato DC3000 (avrRPM1) (or Pst avrRPM1) induces H₂O₂ formation in the chloroplast. Arabidopsis leaves were infiltrated with 1 × 10⁸ UFC/ml Pst avrRPM1 and after 18 h, samples were treated to visualize H₂O₂ formation with the DAB reaction. Controls were infiltrated with 10 mM MgCl₂ and then processed for DAB staining. Then, samples were analyzed in an optical photomicroscope Olympus Provis Model AX70. (B) Effect of FB1 on mitochondria ultrastructure. Wild type Arabidopsis seedlings were treated with FB1 for 72 h and tissues were processed and analyzed according to Saucedo et al.² Ch, chloroplast; M, mitochondria; PM, plasma membrane. Arrows show mitochondrial cisternae. Bars show the correspondent magnification.In addition, we have detected alterations in mitochondria ultrastructure as a result of 72 h of FB1 exposure (Fig. 2B). These alterations mainly consist in the reduced number of cristae, the membrane site of residence of the electron transport complexes. In this sense, it has been shown that factors that induce PCD such as the victorin toxin, methyl jasmonate and H₂O₂ produce alterations in mitochondrial morphology.^20–22 In fact, some of these studies propose that ROS are formed in the mitochondria and then diffuse to the chloroplasts.^22–24It is reasonable to envisage that damage of the membrane integrity of these two organelles reflects the effects of vast amounts of ROS produced by the electron transport chains.^25,26 Recent evidence supports the destruction of the photosynthetic apparatus associated to the generation of ROS in the HR.²⁶ At this time of PCD progression, ROS could be contributing to shut down the energy machinery in the cell, which ultimately would become the point of no-return of PCD²⁷ as part of the execution program of the cell death mediated by LCBs.In conclusion, we propose that ROS can display two different functional roles in the PCD process driven by LCBs. These roles depend on the time of ROS expression, the cellular site where they are generated, the enzymes that produce them, and the magnitude in which they are formed.     

Why have chloroplasts developed a unique motility system?

Organelle movement in plants is dependent on actin filaments with most of the organelles being transported along the actin cables by class XI myosins. Although chloroplast movement is also actin filament-dependent, a potential role of myosin motors in this process is poorly understood. Interestingly, chloroplasts can move in any direction and change the direction within short time periods, suggesting that chloroplasts use the newly formed actin filaments rather than preexisting actin cables. Furthermore, the data on myosin gene knockouts and knockdowns in Arabidopsis and tobacco do not support myosins'' XI role in chloroplast movement. Our recent studies revealed that chloroplast movement and positioning are mediated by the short actin filaments localized at chloroplast periphery (cp-actin filaments) rather than cytoplasmic actin cables. The accumulation of cp-actin filaments depends on kinesin-like proteins, KAC1 and KAC2, as well as on a chloroplast outer membrane protein CHUP1. We propose that plants evolved a myosin XI-independent mechanism of the actin-based chloroplast movement that is distinct from the mechanism used by other organelles.Key words: actin, Arabidopsis, blue light, kinesin, myosin, organelle movement, phototropinOrganelle movement and positioning are pivotal aspects of the intracellular dynamics in most eukaryotes. Although plants are sessile organisms, their organelles are quickly repositioned in response to fluctuating environmental conditions and certain endogenous signals. By and large, plant organelle movements and positioning are dependent on actin filaments, although microtubules play certain accessory roles in organelle dynamics.^1,2 Actin inhibitors effectively retard the movements of mitochondria,^3–6 peroxisomes,^5,7–11 Golgi stacks,^12,13 endoplasmic reticulum (ER),^14,15 and nuclei.^16–18 These organelles are co-aligned and associated with actin filaments.^{5,7,8,10–12,15,18} Recent progress in this field started to reveal the molecular motility system responsible for the organelle transport in plants.¹⁹Chloroplast movement is among the most fascinating models of organelle movement in plants because it is precisely controlled by ambient light conditions.^20,21 Weak light induces chloroplast accumulation response so that chloroplasts can capture photosynthetic light efficiently (Fig. 1A). Strong light induces chloroplast avoidance response to escape from photodamage (Fig. 1B).²² The blue light-induced chloroplast movement is mediated by the blue light receptor phototropin (phot). In some cryptogam plants, the red light-induced chloroplast movement is regulated by a chimeric phytochrome/phototropin photoreceptor neochrome.^23–25 In a model plant Arabidopsis, phot1 and phot2 function redundantly to regulate the accumulation response,²⁶ whereas phot2 alone is essential for the avoidance response.^27,28 Several additional factors regulating chloroplast movement were identified by analyses of Arabidopsis mutants deficient in chloroplast photorelocation.^29–32 In particular, identification of CHUP1 (chloroplast unusual positioning 1) revealed the connection between chloroplasts and actin filaments at the molecular level.²⁹ CHUP1 is a chloroplast outer membrane protein capable of interacting with F-actin, G-actin and profilin in vitro.^29,33,34 The chup1 mutant plants are defective in both the chloroplast movement and chloroplast anchorage to the plasma membrane,^22,29,33 suggesting that CHUP1 plays an important role in linking chloroplasts to the plasma membrane through the actin filaments. However, how chloroplasts move using the actin filaments and whether chloroplast movement utilizes the actin-based motility system similar to other organelle movements remained to be determined.Open in a separate windowFigure 1Schematic distribution patterns of chloroplasts in a palisade cell under different light conditions, weak (A) and strong (B) lights. Shown as a side view of mid-part of the cell and a top view with three different levels (i.e., top, middle and bottom of the cell). The cell was irradiated from the leaf surface shown as arrows. Weak light induces chloroplast accumulation response (A) and strong light induces the avoidance response (B).Here, we review the recent findings pointing to existence of a novel actin-based mechanisms for chloroplast movement and discuss the differences between the mechanism responsible for movement of chloroplasts and other organelles.     

MEK1/2 and p38-like MAP kinase successively mediate H2O2 signaling in Vicia guard cell

As a second messenger, H₂O₂ generation and signal transduction is subtly controlled and involves various signal elements, among which are the members of MAP kinase family. The increasing evidences indicate that both MEK1/2 and p38-like MAP protein kinase mediate ABA-induced H₂O₂ signaling in plant cells. Here we analyze the mechanisms of similarity and difference between MEK1/2 and p38-like MAP protein kinase in mediating ABA-induced H₂O₂ generation, inhibition of inward K⁺ currents, and stomatal closure. These data suggest that activation of MEK1/2 is prior to p38-like protein kinase in Vicia guard cells.Key words: H₂O₂ signaling, ABA, p38-like MAP kinase, MEK1/2, guard cellAn increasing number of literatures elucidate that reactive oxygen species (ROS), especially H₂O₂, is essential to plant growth and development in response to stresses,^1–4 and involves activation of various signaling events, among which are the MAP kinase cascades.^1–3,5 Typically, activation of MEK1/2 mediates NADPH oxidase-dependent ROS generation in response to stresses,^4,6–8 and the facts that MEK1/2 inhibits the expression and activation of antioxidant enzymes reveal how PD98059, the specific inhibitor of MEK1/2, abolishes abscisic acid (ABA)-induced H₂O₂ generation.^6,8,9 It has been indicated that PD98059 does not to intervene on salicylic acid (SA)-stimulated H₂O₂ signaling regardless of SA mimicking ABA in regulating stomatal closure.^2,6,8,10 Generally, activation of MEK1/2 promotes ABA-induced stomatal closure by elevating H₂O₂ generation in conjunction with inactivating anti-oxidases.Moreover, activation of plant p38-like protein kinase, the putative counterpart of yeast or mammalian p38 MAP kinase, has been reported to participate in various stress responses and ROS signaling. It has been well documented that p38 MAP kinase is involved in stress-triggered ROS signaling in yeast or mammalian cells.^11–13 Similar to those of yeast and mammals, many studies showed the activation of p38-like protein kinase in response to stresses in various plants, including Arabidopsis thaliana,^14–16 Pisum sativum,¹⁷ Medicago sativa¹⁸ and tobacco.¹⁹ The specific p38 kinase inhibitor SB203580 was found to modulate physiological processes in plant tissues or cells, such as wheat root cells,²⁰ tobacco tissue²¹ and suspension-cultured Oryza sativa cells.²² Recently, we investigate how activation of p38-like MAP kinase is involved in ABA-induced H₂O₂ signaling in guard cells. Our results show that SB203580 blocks ABA-induced stomatal closure by inhibiting ABA-induced H₂O₂ generation and decreasing K⁺ influx across the plasma membrane of Vicia guard cells, contrasting greatly with its analog SB202474, which has no effect on these events.^23,24 This suggests that ABA integrate activation of p38-like MAP kinase and H₂O₂ signaling to regulate stomatal behavior. In conjunction with SB203580 mimicking PD98059 not to mediate SA-induced H₂O₂ signaling,^23,24 these results generally reveal that the activation of p38-like MAP kinase and MEK1/2 is similar in guard cells.On the other hand, activation of p38-like MAP kinase^23,24 is not always identical to that of MEK1/2^8,25 in ABA-induced H₂O₂ signaling of Vicia guard cells. For example, H₂O₂^- and ABA-induced stomatal closure was partially reversed by SB203580. The maximum inhibition of both regent-induced stomatal closure were observed at 2 h after treatment with SB203580, under which conditions the stomatal apertures were 89% and 70% of the control values, respectively. By contrast, when PD98059 was applied together with ABA or H₂O₂, the effects of both ABA- and H₂O₂-induced stomatal closure were completely abolished (Fig. 1). These data imply that the two members of MAP kinase family are efficient in H₂O₂-stimulated stomatal closure, but p38-like MAP kinase is less susceptive than MEK1/2 to ABA stimuli.Open in a separate windowFigure 1Effects of SB203580 and PD98059 on ABA- and H₂O₂-induced stomatal closure. The experimental procedure and data analysis are according to the previous publication.^8,23,24It has been reported that ABA or NaCl activate p38 MAP kinase in the chloronema cells of the moss Funaria hygrometrica in 2∼10 min.²⁶ Similar to this, SB203580 improves H₂O₂-inhibited inward K⁺ currents after 4 min and leads it to the control level (100%) during the following 8 min (Fig. 2). However, the activation of p38-like MAP kinase in response to ABA need more time, and only recovered to 75% of the control at 8 min of treatment (Fig. 2). These results suggest that control of H₂O₂ signaling is required for the various protein kinases including p38-like MAP kinase and MEK1/2 in guard cells,^1,2,8,23,24 and the ABA and H₂O₂ pathways diverge further downstream in their actions on the K⁺ channels and, thus, on stomatal control. Other differences in action between ABA and H₂O₂ are known. For example, Köhler et al. (2001) reported that H₂O₂ inhibited the K⁺ outward rectifier in guard cells shows that H₂O₂ does not mimic ABA action on guard cell ion channels as it acts on the K⁺ outward rectifier in a manner entirely contrary to that of ABA.²⁷Open in a separate windowFigure 2Effect of SB203580 on ABA- and H₂O₂-inhibited inward K⁺ currents. The experimental procedure and data analysis are according to the previous publication.²⁴ SB203580 directs ABA- and H₂O₂-inactivated inward K⁺ currents across plasma membrane of Vicia guard cells. Here the inward K⁺ currents value is stimulated by −190 mV voltage.Based on the similarity and difference between PD98059 and SB203580 in interceding ABA and H₂O₂ signaling, we speculate the possible mechanism is that the member of MAP kinase family specially regulate signal event in ABA-triggered ROS signaling network,^1–4 and the signaling model as follows (Fig. 3).Open in a separate windowFigure 3Schematic illustration of MAP kinase-mediated H₂O₂ signaling of guard cells. The arrows indicate activation. The line indicates enhancement and the bar denotes inhibition.     

Reconstruction of Metabolic Pathways,Protein Expression,and Homeostasis Machineries across Maize Bundle Sheath and Mesophyll Chloroplasts: Large-Scale Quantitative Proteomics Using the First Maize Genome Assembly

Chloroplasts in differentiated bundle sheath (BS) and mesophyll (M) cells of maize (Zea mays) leaves are specialized to accommodate C₄ photosynthesis. This study provides a reconstruction of how metabolic pathways, protein expression, and homeostasis functions are quantitatively distributed across BS and M chloroplasts. This yielded new insights into cellular specialization. The experimental analysis was based on high-accuracy mass spectrometry, protein quantification by spectral counting, and the first maize genome assembly. A bioinformatics workflow was developed to deal with gene models, protein families, and gene duplications related to the polyploidy of maize; this avoided overidentification of proteins and resulted in more accurate protein quantification. A total of 1,105 proteins were assigned as potential chloroplast proteins, annotated for function, and quantified. Nearly complete coverage of primary carbon, starch, and tetrapyrole metabolism, as well as excellent coverage for fatty acid synthesis, isoprenoid, sulfur, nitrogen, and amino acid metabolism, was obtained. This showed, for example, quantitative and qualitative cell type-specific specialization in starch biosynthesis, arginine synthesis, nitrogen assimilation, and initial steps in sulfur assimilation. An extensive overview of BS and M chloroplast protein expression and homeostasis machineries (more than 200 proteins) demonstrated qualitative and quantitative differences between M and BS chloroplasts and BS-enhanced levels of the specialized chaperones ClpB3 and HSP90 that suggest active remodeling of the BS proteome. The reconstructed pathways are presented as detailed flow diagrams including annotation, relative protein abundance, and cell-specific expression pattern. Protein annotation and identification data, and projection of matched peptides on the protein models, are available online through the Plant Proteome Database.Plants can be classified as C₃ or C₄ species based on the primary product of carbon fixation in photosynthesis. The primary product of carbon fixation is a four-carbon compound (oxaloacetate [OAA]) in C₄ plants but a three-carbon compound (3-phosphoglycerate [3PGA]) in C₃ plants. In leaves of C₄ grasses such as maize (Zea mays), photosynthetic activities are partitioned between two anatomically and biochemically distinct bundle sheath (BS) and mesophyll (M) cells. A single ring of BS cells surrounds the vascular bundle, followed by a concentric ring of specialized M cells, creating the classical Kranz anatomy. Active carbon transport (in the form of C₄ organic acids) from M cell to BS cells and specific expression of Rubisco in the BS cells allows Rubisco, the carboxylating enzyme in the Calvin cycle, to operate in a high CO₂ concentration. The high CO₂ concentration suppresses the oxygenation reaction by Rubisco (and the subsequent energy-wasteful photorespiratory pathway), resulting in increased photosynthetic yield and more efficient use of water and nitrogen. The history of C₄ research has been described (Nelson and Langdale, 1992; Sage and Monson, 1999; Edwards et al., 2001). At present, there is renewed interest in C₄ photosynthesis, stimulated in part by the potential use of C₄ plants as a source of biofuels (Carpita and McCann, 2008) and the genetic engineering of C₄ rice (Oryza sativa; Sheehy et al., 2007; Hibberd et al., 2008; Taniguchi et al., 2008). The use of new genomics and/or proteomics tools has resulted in new insights into cellular differentiation in C₄ plants (Majeran and Van Wijk, 2009).Proteins are responsible for most cellular functions, and knowing their abundance, cell-type specific expression patterns, and subcellular localization is essential to understand C₄ differentiation. Previously, we published a quantitative analysis of purified M and BS chloroplast (soluble) stromal proteomes in which BS-M protein accumulation ratios for 125 accessions were determined; this covered a limited range of plastid functions, although it enabled the integration of information from previous studies (Majeran et al., 2005). A subsequent complementary quantitative proteomics study, using nano-liquid chromatography (LC)-LTQ-Orbitrap mass spectrometry (MS) and label-free spectral counting complemented with other techniques, identified proteins in BS and M thylakoid and envelope membranes of maize chloroplasts and determined cell type-specific differences in (1) the protein assembly state and composition of the four photosynthetic complexes and of a new type of NADPH dehydrogenase (NDH) complex; (2) the auxiliary functions of the thylakoid proteome; and (3) protein and metabolite transport functions of M and BS chloroplast envelopes (Majeran et al., 2008). Comparative MS analysis of chloroplast envelope membranes from leaves of pea (Pisum sativum), a C₃ species, and from M chloroplasts of maize showed an enrichment of several known and putative translocators in the maize M envelopes (Brautigam et al., 2008). The conclusions of these proteome analyses are summarized by Majeran and van Wijk, 2009.Whereas these proteomics studies provide significant progress in understanding the organization of C₄ metabolism in maize, three aspects have not been adequately addressed: (1) the stromal proteomes of BS and M chloroplasts likely each contain more than 1,500 proteins, but the BS-M ratios for only approximately 125 proteins were quantified, resulting in very limited coverage of several important secondary metabolic pathways such as sulfur, fatty acid, amino acid, and nucleotide metabolism; (2) information about relative concentrations of stromal proteins in BS and M chloroplasts is lacking but is needed as a basis for quantitative modeling and metabolic engineering of C₄ photosynthesis and other metabolic pathways; the growing “toolbox” of proteomics and MS now allows for such quantitative analyses (Bantscheff et al., 2007; Kumar and Mann, 2009); (3) the soluble (Majeran et al., 2005) and membrane (Majeran et al., 2008) proteome data sets were analyzed by different techniques and mass spectrometers, mostly due to the improvement of commercial mass spectrometers in that time frame. Therefore, it is difficult to understand the quantitative relationships between these data sets. This study addresses these three aspects.So far, maize proteome analyses used essentially ZmGI maize assemblies (for the Z. mays Gene Index) based on ESTs, combined with a limited amount of additional DNA sequence information. The ZmGI was originally generated by The Institute for Genome Research and subsequently supported by the computational Biology and Functional Genomics Laboratory (http://compbio.dfci.harvard.edu/index.html). This ZmGI database did not have annotated gene models (for proteome analysis, the DNA sequences were searched in all six reading frames), and low expressed genes were likely underrepresented. In our most recent BS-M chloroplast analyses (Majeran et al., 2008) as well as a maize envelope analysis (Brautigam et al., 2008), the MS data were searched against ZmGI version 16.0 or 17.0. Since that time, the maize genome has been sequenced (using a bacterial artificial chromosome approach), a physical map was created (the maize accessioned golden path AGP version 1), and its first assembly with gene coordinates and predicted proteins was very recently released (June 2009; http://ftp.maizesequence.org/release-4a.53/sequences/) and published (Schnable et al., 2009). This release contains 32,540 genes with 53,764 gene models; most of the gene models are evidence based. The new maize genome assembly is expected to improve maize proteome analysis with more accurate protein identification and quantitative assessment of protein expression patterns. This also allows for the determination of N-terminal localization signals, which was rarely possible from EST assemblies, as N termini were often lacking.This study presents a quantitative protein expression atlas of differentiated maize leaf M and BS chloroplasts using high-resolution and mass-accuracy MS (using a LTQ-Orbitrap) and the new maize genome assembly. Three biological replicates of stromal proteomes of isolated BS and M chloroplasts were analyzed. Quantification was carried out based on the “spectral counting” method (Zybailov et al., 2005, 2008; Bantscheff et al., 2007; Choi et al., 2008) using a sophisticated bioinformatics “workflow” in particular to deal with gene duplications and extended gene families observed in polyploids such as maize. These new stromal data sets were combined with a reanalysis of our recent BS and M membrane proteome data sets (Majeran et al., 2008) against genome 4a53. Compared with previous maize leaf proteome analyses, this study provides an integrated overview of both primary and especially secondary metabolism, as well as chloroplast gene expression and protein biogenesis, in far greater depth. The reconstructed pathways are presented as figures that include quantitative protein information; pathways include primary carbon metabolism, starch metabolism, nucleotide metabolism, fatty acid and lipid biosynthesis, chlorophyll, heme, and carotenoid synthesis, and nitrogen assimilation. We briefly comment on the use of the new maize genome assembly for proteome analysis. All matched peptides are projected on the predicted protein models via the Plant Proteomics Database (PPDB; http://ppdb.tc.cornell.edu/). Interactive functional annotation, chloroplast localization assignments, as well as details of protein identification are also available via PPDB.     

ROS removal by DJ-1: Arabidopsis as a new model to understand Parkinson disease

Reactive oxygen species represent one of the principal factors that cause cell death and scavenging of reactive oxygen species by superoxide dismutase-related pathway is essential for cell survival. The Parkinson disease-related DJ-1 protein (also known as PARK7) has been implicated in resistance against oxidative stress in dopaminergic neurons however, its molecular mechanism has to date been unknown. We have used Arabidopsis thaliana as a model system to demonstrate that DJ-1, in both plant and mammalian cells, directly influence SOD activity in a highly conserved manner thereby preventing cell death. These data not only provides evidence for the molecular mechanisms associated with DJ-1-induced Parkinson disease but also highlight the unprecedented value of plants as a tool in understanding human disease mechanisms.Key words: DJ-1, stress, cell death, Parkinson disease, ArabidopsisReactive oxygen species (ROS) are involved in a myriad of fundamental biological processes including cell signaling and cellular defense pathways in plants and animals.^1–3 Despite its role as a signaling molecule, inappropriate and elevated levels of ROS have a major impact on the etiology of neurodegenerative diseases, such as for example Parkinson disease (PD), and in oxidative stress responses in plants. In general ROS can cause damage to DNA, lipids, proteins and various cofactors. During normal physiological conditions, when ROS are continuously generated, antioxidant defense systems are adequately equipped to prevent ROS-induced tissue dysfunction.^4,5 However upon elevated ROS generation the cellular antioxidant systems either recruit additional factors to minimize ROS-induced damage or cells suffer the consequences of cell death. Because of this dichotomy, where ROS plays a vital role during growth and development but can also have overwhelming damaging effects, it is clear that strict regulatory mechanisms need to be in place to effectively control ROS levels. In both plant and mammalian cells elevated ROS levels lead to cell death and in various human disease such as PD, Alzheimer disease, amyotrophic lateral sclerosis and Huntington disease proteins involved in stress-related pathways are often mutated.⁶In both plants and mammals mitochondria act as an important source of ROS however, plants also produce ROS in chloroplasts as part of photosynthetic activity. Combined with the fact that plants are sessile organisms it suggests that, although similar in nature, plants most probably have more complex antioxidant systems than other organisms.Strategies for removing excess ROS are similar in plants and humans. The principle ROS removal pathway involves superoxide dismutases (SOD) (or copper/zinc superoxide dismutase-CSD in plants), glutathione peroxidases (GPX) and catalases (CAT) localized in the cytosol, mitochondria and chloroplasts (Fig. 1). SOD converts superoxide anion to H₂O₂, which is then detoxified to H₂O by GPX and CAT. In Arabidopsis, besides SOD, GPX and CAT, there are five ascorbate peroxidases (APX) located in the cytosol and chloroplasts, involved in scavenging ROS generated during photosynthesis.^7,8 Therefore, SOD, GPX, CAT and APX, together with other auxiliary proteins, form the main line of defense against ROS.Open in a separate windowFigure 1Involvement of DJ-1-like proteins in ROS scavenging pathways. Produced O₂^- is converted to H₂O₂ by SOD in human or CSD (Arabidopsis SOD) in Arabidopsis. H₂O₂ is then converted to H₂O by GPX or CAT (catalase) in human or APX, GPX or CAT in Arabidopsis. DJ-1-like proteins interact with SOD or GPX in humans and CSD, GPX and APX in Arabidopsis. It is assumed here that DJ-1-like proteins may also interact with catalases (broken arrows).DJ-1 was originally identified as an oncogene and represents a ubiquitous redox-responsive cytoprotective protein with diverse functions where one of its main roles have been attributed to oxidative stress protection.⁹ Numerous studies have shown that several DJ-1 mutations in humans cause autosomal recessive, early onset PD however, its mode of action has been elusive in terms of having a direct influence on neuronal cell death.¹⁰ In an attempt to clarify the mechanism of DJ-1 we established Arabidopsis thaliana as a new and novel model system.¹¹ The Arabidopsis genome contains three DJ-1 homologs compared to the single DJ-1 locus found in humans and we showed that two of these (AtDJ-1b and AtDJ-1c) localizes to chloroplasts whilst one, AtDJ-1a, localizes to the cytosol and nucleus as observed for human DJ-1.¹¹ As mutated DJ-1 in mammals leads to cell death we identified and characterized a DJ-1 loss-of-function mutant which showed increased cell death in aging plants. Using Bimolecular Fluorescence Complementation (BiFC) and isothermal titration calorimetry (ITC) assays we showed that AtDJ-1a interacts with CSD1, the cytosolic SOD in Arabidopsis, and with human SOD1 in plant cells. Further we demonstrated that the human DJ-1 protein interacts with SOD1 in mammalian CHO cells.¹¹ Similar approaches were also employed to show that AtDJ-1a and human DJ-1 had an interaction with GPX2 in plant and mammalian cells.¹¹Enzyme assays revealed that AtDJ-1a and DJ-1 stimulated SOD/CSD1 activity and that only the copper-loaded forms of AtDJ-1a and DJ-1 had this effect suggesting that AtDJ-1a/DJ-1 may provide copper for SOD/CSD1.¹¹ Although the observed SOD activation provides clues towards the role of DJ-1 in detoxification of ROS, SOD only converts superoxide anion to H₂O₂ which must further be detoxified to H₂O by GPX and CAT. Although we showed that AtDJ-1a and human DJ-1 can interact with AtGPX2 and GPX2, respectively, we observed no changes in GPX2 activity upon DJ-1 interaction. The reason for this may be several-fold. First, cellular GPX2 activity levels may be sufficient to convert SOD-generated H₂O₂ to H₂O. Second, DJ-1 may indeed have no effect on GPX2 activity but simply act as an anchor to dock GPX2 in the vicinity of SOD. To test whether the DJ-1/SOD/GPX2 complex recruits other auxiliary proteins we have also shown that AtDJ-1a interacts with the Arabidopsis cytosolic APX1 protein (Fig. 2, unpublished data). It is also highly possible that DJ-1 interacts with catalase or at least influences its activity (Fig. 1). Although we have no data to date indicating a functional significance of the DJ-1/APX1 interaction we speculate that DJ-1 indeed acts as a scaffold protein bringing together SOD, GPX and possibly APX1 to mediate and control ROS scavenging, ultimately preventing oxidative stress-induced cell death (Fig. 3).Open in a separate windowFigure 2Interaction of AtDJ-1a with APX1. AtDJ-1a tagged with the N-terminal region of GFP and APX1 tagged with the C-terminal region of GFP gene were co-transformed into tobacco cells. The observed GFP signal in (B) demonstrates an AtDJ-1a/APX1 interaction through reconstitution of functional GFP molecules. (A) Negative control.Open in a separate windowFigure 3Working model of AtDJ-1a and DJ-1 mode of action. AtDJ-1a and DJ-1 interacts with SOD and GPX2 leading to SOD activation in a copper-dependent fashion. It is proposed that AtDJ-1a and DJ-1 delivers copper to SOD enhancing its activity whilst GPX2 is anchored by AtDJ-1 and DJ-1 to the protein complex to ensure conversion of the SOD-generated H₂O₂ to H₂O.The fact that Arabidopsis has three DJ-1 homologs where two of these, AtDJ-1b and AtDJ-1c, are localized to chloroplasts¹¹ underlines the protective role of DJ-1-like proteins during oxidative stress in plants. From our localization studies it appears that AtDJ-1b is localized to the chloroplast stroma whilst AtDJ-1c is localized to both the stroma and the thylakoid membranes (unpublished data). Whether AtDJ-1b and AtDJ-1c act in isolation or in concert and how these two proteins are involved in photosynthesis-induced ROS regulation is unclear but represent exciting future challenges.The notion that plants can be used as tools to increase our understanding of human disease mechanisms is somewhat obscure to the general scientific community. The fact remains that many discoveries with direct relevance to human health and disease have been elaborated using Arabidopsis, and several processes important to human biology are more easily studied in this versatile model plant.¹² The use of Arabidopsis to understand human disease states has several advantages: (1) Arabidopsis represents a well established model organism with a fully annotated genome, (2) The Arabidopsis genome contains homologs of numerous genes involved in human disease, (3) The identification and generation of Arabidopsis mutants is simple and requires little effort, (4) Arabidopsis growth and maintenance requires little infrastructure and running costs and (5) Arabidopsis research has few ethical constraints.Despite the advantages of Arabidopsis as a model system for elucidating human disease mechanisms it is important to appreciate that Arabidopsis and plant research in general can only reach its full potential in the field of medical research if combined with complementary, and perhaps more conventional, model systems.     

Tubular actin filaments in tobacco guard cells

The dynamic remodeling of actin filaments in guard cells functions in stomatal movement regulation. In our previous study, we found that the stochastic dynamics of guard cell actin filaments play a role in chloroplast movement during stomatal movement. In our present study, we further found that tubular actin filaments were present in tobacco guard cells that express GFP-mouse talin; approximately 2.3 tubular structures per cell with a diameter and height in the range of 1–3 µm and 3–5 µm, respectively. Most of the tubular structures were found to be localized in the cytoplasm near the inner walls of the guard cells. Moreover, the tubular actin filaments altered their localization slowly in the guard cells of static stoma, but showed obvious remodeling, such as breakdown and re-formation, in moving guard cells. Tubular actin filaments were further found to be colocalized with the chloroplasts in guard cells, but their roles in stomatal movement regulation requires further investigation.Key words: actin dynamics, tubular actin filaments, chloroplast, guard cell, stomatal movementStomatal movement responses to surrounding environment are mediated by guard cell signaling.^1,2 Actin filaments within guard cells are dynamic cytoarchitectures and function in stomatal development and movement.³ Arrays of actin filaments in guard cells that are dependent on different stomatal apertures have also been reported in references ^4–7. For example, the random or longitudinal orientations of actin filaments in closed stomata change to a radial orientation or ring-like array after stomata opening.^5,6,8 The reorganization of the actin architecture during stomatal movement depends on the depolymerization and repolymerization of actin filaments in guard cells. In contrast to the traditional treadmill model of actin dynamic mechanisms, stochastic dynamics of actin have been revealed in plant cells, such as in the epidermal cells of hypocotyl and root, the pavement cells of Arabidopsis cotyledons, and the guard cells of tobacco (Nicotiana tabacum).^9–11 In this alternative system, the short actin fragments generated from severed long filaments can link with each other to form longer filaments by end-joining activity. The actin regulatory proteins, Arp2/3 complex, capping protein and actin depolymerizing factor (ADF)/cofilin, may also be involved in the stochastic dynamics of actin filaments.^12,13Using tobacco GFP-mouse talin expression lines, we have previously analyzed the stochastic dynamics of guard cell actin filaments and their roles in chloroplast displacement during stomatal movement.^6,11 We found from these analyses that another arrangement of actin filaments, i.e., tubular actin filaments, exists in the guard cells of these tobacco lines. We first found the circle-like actin filaments in 82% of the guard cells (counting 320 cells) in tobacco expressing GFPmouse talin when analyzing a single optical section (Fig. 1A). In a previous study of BY-2 cells expressing GFP-Lifeact labeled actin filaments, Smertenko et al. found similar structures, i.e., quoit-like structures or acquosomes in all of the plant tissues examined except growing root hairs.¹⁰ However, in our present analysis of serial sections, we determined that the circle-like actin filaments in the tobacco guard cells were long tubes (Fig. 1A), as the lengths (about 3–5 µm) of these structures were greater than their diameter (about 1–3 µm). Hence, we denoted these structures as tubular actin filaments to distinguish them from the circular conformations of actin filaments observed previously in other plant cell tissues.^10,14–19 About 2.3 of these tubular actin filaments were found per guard cell, which is less than the number of acquosomes reported in BY-2 cells (about 6.7 per cell).¹⁰ Analysis of serial optical sections at the z-axis revealed that the tubular actin filaments localize in the cytoplasm near the inner walls of the guard cells (Fig. 1B), which is similar to the distribution of chloroplasts in guard cells.¹¹ Longitudinal sections further revealed a colocalization of tubular actin filaments and chloroplasts (Fig. 1B).Open in a separate windowFigure 1Tubular actin filaments in the guard cells of a tobacco (Nicotiana tabacum) line expressing GFP-mouse talin. (A) Optical-sections (interval, 1.5 µm) of guard cells in a moving stoma showing tubular actin filaments (arrow heads). Frames (a1) and (a2) are cross sections of 1.5-µm-picture through the yellow and red lines, respectively, revealing the cross section of the circle structures are parallel lines (arrows). (B) Optical-sections of a stoma from the outer periclinal walls to the inner walls of the guard cells (interval, 1 µm). The tubular actin filaments (arrow heads) are localized in the cytoplasm near to the inner periclinal walls of guard cells. Frame (b1) is the guard cell on the right of the frame “4 µm”; (b2) is the cross section of b1 through the red line; and (b3) is a higher magnification image of the area encompassed by the white square in b2. Arrows indicate the colocalization between the tubular actin filaments and the chloroplast (indicated using a red pseudocolor). (C) Time-series imaging showing the movement of tubular actin filaments in the guard cells of static stomata. Frame (c1) comprises three images colored red (0 S), green (40 S) and blue (80 S), that are merged in a single frame to show the translocation of the tubular actin filaments (arrows). (D) Time-series images of the opening stomata showing the breakdown (arrows) and re-formation (arrowheads) of the tubular actin filaments. All images were captured using a Zeiss LSM 510 META confocal laser scanning microscope, as described by Wang et al.¹¹ Bars, 10 µm.We performed time-lapse imaging and found that the translocation of tubular actin filaments is slow in static stomata in which the distance between two tubular actin filaments typically increased from 2.22 to 2.50 µm after 80 sec (Fig. 1C). In moving stomata, however, the tubular actin filaments showed an obvious dynamic reorganization whereby they could be processed into short fragments and also reemerged after they had disintegrated (Fig. 1D). These results indicate that tubular actin filaments have stochastic dynamics that are similar to the long actin filaments of guard cells.¹¹ In our previous study, we found that the stochastic dynamics of actin filaments correlate with light-induced chloroplast movement in guard cells.¹¹ However, whether the dynamics of the tubular actin filaments are also involved in chloroplast movement during stomatal movement remains to be investigated. In cultured mesophyll cells which had been mechanically isolated from Zinnia elegans, Wilsen et al. previously found a close association between fully closed actin rings and chloroplasts.¹⁸ These authors further found that the average percentage of cells with free actin rings increased at the initial culture stage, and then decreased, which indicates that the formation of actin rings might be a response of the actin cytoskeleton to cellular stress or disturbance.¹⁸ The turgor pressure of guard cells is the fundamental basis of stomatal movement leading to changes in the shape, volume, wall structure, and membrane surface of guard cells.^20–24 We speculate from our current data that there is a relationship between tubular actin filaments and the shape changes of guard cells during stomatal movement.     

Formation of xanthine and the use of purine metabolites as a nitrogen source in Arabidopsis plants

In our recent paper in The Plant Journal,¹ we described the remobilization of purine metabolites during natural and dark induced senescence in wild type and Atxdh1 mutant lines impaired in xanthine dehydrogenase (XDH), a pivotal enzyme in the purine catabolism pathway. In the light of these observations and additional evidence shown here, we discuss the probable pathways leading to xanthine synthesis in Arabidopsis plants during senescence and the role that purine metabolites play as an ongoing source of nitrogen in plant growth.Key words: hypoxanthine, purine catabolism, senescence, xanthine, xanthine dehydrogenaseIn mammalian purine catabolism, hypoxanthine is oxidized to xanthine by xanthine oxidase.² In planta, xanthine can be synthesized in the purine degradation pathway, via three alternative precursors, guanine, xanthosine or hypoxanthine³ (Fig. 1A). Thus, the exact pathway leading to xanthine may depend on the species examined, the particular plant organ, developmental stage or specific environmental stimuli. For example, guanine and guanosine were shown to be the main precursor of ureides and CO₂ in cacao leaves⁴ while in tea leaves, elevated amounts of labeled xanthosine were recovered as ureides.^5,6 However, when hypoxanthine was used as a substrate for inosine monophosphate (IMP) formation in tobacco protoplasts⁷ more than 90% of labeled hypoxanthine was recovered as salvage products, nucleotides and RNA and only less then 10% was found as ureides in cacao leaves.⁴ Furthermore, when [8-¹⁴C]-hypoxanthine is supplied to soybean embryo axes or Jerusalem artichoke shoots it selectively labelled the guanine nucleotide pool.^3,8,9 These data do not support the possibility of hypoxanthine being a direct precursor for xanthine formation and illustrate the concept of species dependent differences in xanthine biosynthesis.¹⁰Open in a separate windowFigure 1Purine catabolism, xanthine and hypoxanthine accumulation and Arabidopsis plants growth. (A) Purine nucleotide catabolism in plants. Enzymes shown are: (1) AMP deaminase (EC 3.5.4.6), (2) IMP dehydrogenase (EC 1.1.1.205), (3) GMP synthase (EC 6.3.5.2), (4) 5''-nucleotidase, (5) Nucloeside phosphotransferase (EC 2.7.1.77), (6) Inosine-guanosine nucleosidase (EC 3.2.2.2), (7) Guanine deaminase (EC 3.5.4.15), (8) Xanthine dehydrogenase (EC 1.1.1.204), (9) Uricase EC 1.7.3.3, (10) Hydroxyisourate hydrolase (EC 3.5.2.17),1,18 (11) Allantoinase, allantoin amidohydrolase (EC 3.5.2.5), (12) Allantoicase, allantoate amidohydrolase (EC 3.5.3.4), (13) Ureidoglycolate lyase (EC 4.3.2.3), (14) Urease EC 3.5.1.5, (15) Allantoin deaminase (EC 3.5.3.9), (16) Ureidoglycine amidohydrolase (EC 3.5.3.-), (17) Ureidoglycolate hydrolase (EC 3.5.3.19). (B) Analysis of the purine metabolites, hypoxanthine and xanthine, in response to dark stress. Hypoxanthine and xanthine were determined by HPLC¹ in rosette leaves of wild-type (Col) and Ri14, XDH1 RNA interference plants after being kept in dark for 6 days and transferred to a 16-h light/8-h dark regime for recovery over an additional 3 days. Values are means ± SEM (n = 3). (C) Wild-type (Col) and XDH-compromised plants (KO, SALK_148364; Ri, XDH1 RNA interference) were germinated on ¼ MS medium and transplanted on the 5 day to a full MS medium (upper panel) or MS medium with 5.0 mM xanthine and urea as the sole nitrogen source. After transplanting the seedlings were left to grow for 14 days under a 16-h light/8-h dark regime (100 µmol m⁻² sec⁻¹) and then photographed. Leaf size was estimated using ImageJ software (http://rsb.info.nih.gov/ij/). Values are means ± SEM (n = 3).To study the possible role of hypoxanthine in xanthine formation in Arabidopsis we utilized XDH1 mutants. The mutants do not show any detectable XDH activity in-vitro when using hypoxanthine and/or xanthine as substrates.^1,11 Furthermore, no other enzyme is known to catalyze the conversion of hypoxanthine to xanthine, other than the molybdenum cofactor containing-XDH1. Yet, xanthine accumulation was readily detected in mutant leaves and was up to 100-fold higher than hypoxanthine either in normal growth conditions (Fig. 1B, time 0) or when exposed to dark induced senescence and to a light recovery period thereafter (Fig. 1B). These results indicate that most likely, hypoxanthine is not a major direct source for xanthine formation in Arabidopsis. The results imply that xanthosine or guanine are a source, although, one cannot exclude the possibility that hypoxanthine could be converted to xanthine in a pathway leading to inosine, IMP and then either via guanine or xanthosine, back to xanthine as illustrated in Figure 1A.In legumes inoculated with rhizobia, nitrogen is fixed initially as NH₃/NH₄⁺ that is subsequently incorporated through the purine pathway to form IMP, and finally ureides. The central role of purine catabolism in plant nitrogen metabolism was demonstrated mainly in legumes in which the purine nucleotides are degraded via uric acid and allantoin to urea and then to CO₂ and NH₃, which is then re-assimilated via the glutamine oxoglutarate aminotransferase (GOGAT) pathway (reviewed in ref. ³). What then is the role of purine catabolism pathways in non leguminous plants? Are the nitrogenous products of the degraded purines re-assimilated in non-legumes as in legume plants? We recently showed in Arabidopsis that a marked transition from assimilation, during the plants normal growth, to a state of rapid metabolite turnover occurs when plants were exposed to extended dark stress, senescence or even during normal diurnal cycles.¹⁰ This was depicted by the acceleration of purine catabolic recycling activities in which XDH1 plays a central role.¹ To test for a possible role of the accumulating purines as a source of nitrogen metabolites, we grew wild-type Arabidopsis plants and their XDH1 mutants under heterotrophic conditions. The agar plates contained either full MS nutrient solution with nitrate and ammonia or the purine metabolites, hypoxanthine (data not shown), xanthine or urea (Fig. 1C) as sole nitrogen source. The results show that the mutant plants exhibited slower growth in the medium contained xanthine or hypoxanthine compared to wild-type (Fig. 1C, lowest insert). The suboptimal growth of wild type lines is likely due to the low solubility of hypoxanthine and xanthine. In contrast, the growth on urea was the same for wild-type and XDH1 mutant transgenic plants (Fig. 1C). These results suggest that the conversion of xanthine to metabolically active intermediates, such as ureides and urea synthesized through XDH1, can play a role in ensuring nutrient supply for normal plant growth in purine containing media. Indeed, urea has been shown to be essential for the germination of Arabidopsis under nitrogenlimited conditions,¹² and recent studies have also shown that uric acid,¹³ allantoin and allantoate,^14–16 can serve as the sole nitrogen source during the growth of Arabidopsis plants. Taken together, the data suggest that ureide formation is an active component of normal plant metabolism facilitating the recovery of nitrogen in stress and non-stressed metabolism in a manner analogous to legumes. Indeed, legumes arose about 50–55 milion years ago¹⁷ and likely recruited and amplified existing plant functional purine pathways for their efficient nitrogen distribution system.     

Long-distance trafficking of macromolecules in the phloem

The fact that macromolecules such as proteins and mRNAs overcome the symplastic barriers between various tissue domains was first evidenced by the movement of plant viruses. We have recently demonstrated that viral infection disengages the symplastic restriction present between the sieve element-companion cell complex and neighboring cells in tobacco plants. As a result, green fluorescent protein, which was produced in mesophyll and bundle sheath cells, could traffic into the sieve tube and travel long distances within the vascular system. In this addendum we discuss the likely existence of a novel plant communication network in which macromolecules also act as long-distance trafficking signals. Plasmodesmata interconnecting sieve elements and companion cells as well as plasmodesmata connecting the sieve tube with neighboring cells may play a central role in establishing this communication network.Key words: companion cells, cucumber mosaic virus, Cucumis melo, plasmodesmata, movement protein, sieve-elementsTranslocation of photoassimilates from the source (site of synthesis) to various sink organs is governed, in part, by short-distance intercellular transfer of assimilates to the loading region of the phloem and long-distance transport within the plant vascular system. Sucrose, which is synthesized in the leaf mesophyll, moves cell-to-cell symplastically through plasmodesmata until it reaches the boundary of the sieve element (SE)-companion cell (CC) complex. In many plant species, the connection between phloem parenchyma (PP)/bundle sheath (BS) cells and CCs is characterized by a sparseness of plasmodesmata (e.g., Solanaceae), and sucrose is exported out of the cells to the apoplast. This type of plants (apoplastic loaders) uses sucrose proton symporters to load the sucrose into the vasculature.¹ Cucurbits are considered one of the model plants for symplastic phloem loading.² This type of plant is characterized by abundant plasmodesmata interconnecting the intermediary cells, which are specialized CCs, with the neighboring BS cells. It is generally accepted that in these plants, phloem loading includes intercellular movement of sucrose through the plasmodesmata, along the entire pathway from the mesophyll cell to the SE-CC complex.Interestingly, the existence of plasmodesmata interconnecting the SE-CC complex and neighboring cells is evident in all plant species that are characterized by an apoplastic phloem-loading mechanism. Moreover, microinjection experiments have indicated that plasmodesmata interconnecting the PP-CC are functional, in that they allow the exchange of small membrane-impermeable fluorescent probes.³ Virus movement through plasmodesmata from the mesophyll into the SEs further supports the notion that the symplastic communication between the CC-SE complex and the neighboring cells is functional.⁴One can assume that in apoplastic-loading plants, it would be an advantage to maintain the SE-CC complex as an isolated domain, with no functional plasmodesmata interconnecting it to the neighboring tissue. Symplastic continuity between the two domains could result in leakage of sucrose out of the vasculature and a significant reduction in the efficacy of sucrose loading. The fact that the two domains are interconnected suggests that any back-leakage of sucrose that might occur is insignificant relative to the likely efficacy of this communication route.What might the advantage be for symplastic communication between the SE-CC complex and the neighboring tissue? Accumulated evidence suggests that at the tissue/organ level, cell-to-cell trafficking of information molecules allows for noncell-autonomous control over a range of processes, whereas at the organismal level, the phloem serves as an information superhighway, delivering a wide range of macromolecules to enable the plant to function as a whole organism.^5–8 We advanced the hypothesis that plasmodesmata interconnecting the CCs and PP/BS cells play a pivotal role in controlling the long-distance trafficking of putative signaling molecules.     

ARF-GTPase as a Molecular Switch for Polar Auxin Transport Mediated by Vesicle Trafficking in Root Development

Polar auxin transport (PAT), which is controlled precisely by both auxin efflux and influx facilitators and mediated by the cell trafficking system, modulates organogenesis, development and root gravitropism. ADP-ribosylation factor (ARF)-GTPase protein is catalyzed to switch to the GTP-bound type by a guanine nucleotide exchange factor (GEF) and promoted for hybridization to the GDP-bound type by a GTPase-activating protein (GAP). Previous studies showed that auxin efflux facilitators such as PIN1 are regulated by GNOM, an ARF-GEF, in Arabidopsis. In the November issue of The Plant Journal, we reported that the auxin influx facilitator AUX1 was regulated by ARF-GAP via the vesicle trafficking system.¹ In this addendum, we report that overexpression of OsAGAP leads to enhanced root gravitropism and propose a new model of PAT regulation: a loop mechanism between ARF-GAP and GEF mediated by vesicle trafficking to regulate PAT at influx and efflux facilitators, thus controlling root development in plants.Key Words: ADP-ribosylation factor (ARF), ARF-GAP, ARF-GEF, auxin, GNOM, polar transport of auxinPolar auxin transport (PAT) is a unique process in plants. It results in alteration of auxin level, which controls organogenesis and development and a series of physiological processes, such as vascular differentiation, apical dominance, and tropic growth.² Genetic and physiological studies identified that PAT depends on efflux facilitators such as PIN family proteins and influx facilitators such as AUX1 in Arabidopsis.Eight PIN family proteins, AtPIN1 to AtPIN8, exist in Arabidopsis. AtPIN1 is located at the basal side of the plasma membrane in vascular tissues but is weak in cortical tissues, which supports the hypothesis of chemical pervasion.³ AtPIN2 is localized at the apical side of epidermal cells and basally in cortical cells.^1,4 GNOM, an ARF GEF, modulates the localization of PIN1 and vesicle trafficking and affects root development.^5,6 The PIN auxin-efflux facilitator network controls root growth and patterning in Arabidopsis.⁴ As well, asymmetric localization of AUX1 occurs in the root cells of Arabidopsis plants,⁷ and overexpression of OsAGAP interferes with localization of AUX1.¹ Our data support that ARF-GAP mediates auxin influx and auxin-dependent root growth and patterning, which involves vesicle trafficking.¹ Here we show that OsAGAP overexpression leads to enhanced gravitropic response in transgenic rice plants. We propose a model whereby ARF GTPase is a molecular switch to control PAT and root growth and development.Overexpression of OsAGAP led to reduced growth in primary or adventitious roots of rice as compared with wild-type rice.¹ Gravitropism assay revealed transgenic rice overxpressing OsAGAP with a faster response to gravity than the wild type during 24-h treatment. However, 1-naphthyl acetic acid (NAA) treatment promoted the gravitropic response of the wild type, with no difference in response between the OsAGAP transgenic plants and the wild type plants (Fig. 1). The phenotype of enhanced gravitropic response in the transgenic plants was similar to that in the mutants atmdr1-100 and atmdr1-100/atpgp1-100 related to Arabidopsis ABC (ATP-binding cassette) transporter and defective in PAT.⁸ The physiological data, as well as data on localization of auxin transport facilitators, support ARF-GAP modulating PAT via regulating the location of the auxin influx facilitator AUX1.¹ So the alteration in gravitropic response in the OsAGAP transgenic plants was explained by a defect in PAT.Open in a separate windowFigure 1Gravitropism of OsAGAP overexpressing transgenic rice roots and response to 1-naphthyl acetic acid (NAA). (A) Gravitropism phenotype of wild type (WT) and OsAGAP overexpressing roots at 6 hr gravi-stimulation (top panel) and 0 hr as a treatment control (bottom panel). (B) Time course of gravitropic response in transgenic roots. (C and D) results correspond to those in (A and B), except for treatment with NAA (5 × 10⁻⁷ M).The polarity of auxin transport is controlled by the asymmetric distribution of auxin transport proteins, efflux facilitators and influx carriers. ARF GTPase is a key member in vesicle trafficking system and modulates cell polarity and PAT in plants. Thus, ARF-GDP or GTP bound with GEF or GAP determines the ARF function on auxin efflux facilitators (such as PIN1) or influx ones (such as AUX1).ARF1, targeting ROP2 and PIN2, affects epidermal cell polarity.⁹ GNOM is involved in the regulation of PIN1 asymmetric localization in cells and its related function in organogenesis and development.⁶ Although VAN3, an ARF-GAP in Arabidopsis, is located in a subpopulation of the trans-Golgi transport network (TGN), which is involved in leaf vascular network formation, it does not affect PAT.¹⁰ OsAGAP possesses an ARF GTPase-activating function in rice.¹¹ Specifically, our evidence supports that ARF-GAP bound with ARF-GTP modulates PAT and gravitropism via AUX1, mediated by vesicle trafficking, including the Golgi stack.¹Therefore, we propose a loop mechanism between ARF-GAP and GEF mediated by the vascular trafficking system in regulating PAT at influx and efflux facilitators, which controls root development and gravitropism in plants (Fig. 2). Here we emphasize that ARF-GEF catalyzes a conversion of ARF-bound GDP to GTP, which is necessary for the efficient delivery of the vesicle to the target membrane.¹² An opposite process of ARF-bound GDP to GTP is promoted by ARF-GTPase-activating protein via binding. A loop status of ARF-GTP and ARF-GDP bound with their appurtenances controls different auxin facilitators and regulates root development and gravitropism.Open in a separate windowFigure 2Model for ARF GTPase as a molecular switch for the polar auxin transport mediated by the vesicle traffic system.     

The Arabidopsis peroxisome division mutant pdd2 is defective in the DYNAMIN-RELATED PROTEIN3A (DRP3A) gene

In plants, the division of peroxisomes is mediated by several classes of proteins, including PEROXIN11 (PEX11), FISSION1 (FIS1) and DYNAMIN-RELATED PROTEIN3 (DRP3). DRP3A and DRP3B are two homologous dynamin-related proteins playing overlapping roles in the division of both peroxisomes and mitochondria, with DRP3A performing a stronger function than DRP3B in peroxisomal fission. Here, we report the identification and characterization of the peroxisome division defective 2 (pdd2) mutant, which was later proven to be another drp3A allele. The pdd2 mutant generates a truncated DRP3A protein and exhibits pale green and retarded growth phenotypes. Intriguingly, this mutant displays much stronger peroxisome division deficiency in root cells than in leaf mesophyll cells. Our data suggest that the partial GTPase effector domain retained in pdd2 may have contributed to the distinct mutant phenotype of this mutant.Key words: peroxisome division, dynamin-related protein, arabidopsisIn eukaryotic cells, peroxisomes are surrounded by single membranes and house a variety of oxidative metabolic pathways such as lipid metabolism, detoxification and plant photorespiration.^1,2 To accomplish multiple tasks, the morphology, abundance and positioning of peroxisomes need to be highly regulated. Three families of proteins, whose homologs are present across different kingdoms, have been shown to be involved in peroxisome division in Arabidopsis. The PEX11 protein family is composed of five integral membrane proteins with primary roles in peroxisome elongation/tubulation, the initial step in peroxisome division.^3–5 Although the exact function of PEX11s has not been demonstrated, these proteins are believed to participate in peroxisome membrane modification.^6,7 The FIS1 family consists of two isoforms, which are C-terminal tail-anchored membrane proteins with rate limiting functions at the fission step.^8,9 DRP3A and DRP3B belong to a superfamily of dynamin-related proteins, which are large and self-assembling GTPases involved in the fission and fusion of membranes by acting as mechanochemical enzymes or signaling GTPases.¹⁰ The function of PEX11 seems to be exclusive to peroxisomes, whereas DRP3 and FIS1 are shared by the division machineries of both peroxisomes and mitochondria in Arabidopsis.^8,9,11–16 FIS1 proteins are believed to tether DRP proteins to the peroxisomal membrane,^17,18 but direct evidence has not been obtained from plants. DRP3A and DRP3B share 77% sequence identity at the protein level and are functionally redundant in regulating mitochondrial division; however, DRP3A''s role on the peroxisome seems stronger and cannot be substituted by DRP3B in peroxisome division.^8,13,15In a continuous effort to identify components of the plant peroxisome division apparatus from Arabidopsis, we performed genetic screens in a peroxisomal marker background expressing the YFP (yellow fluorescent protein)-PTS1 (peroxisome targeting signal 1, containing Ser-Lys-Leu) fusion protein. Mutants with defects in the morphology and abundance of fluorescently labeled peroxisomes are characterized. Following our analysis of the pdd1 mutant, which turned out to be a strong allele of DRP3A,⁸ we characterized the pdd2 mutant.In root cells of the pdd2 mutant, extremely elongated peroxisomes and a beads-on-a-string peroxisomal phenotype are frequently observed (Fig. 1A and B). These peroxisome phenotypes resemble those of pdd1 and other strong drp3A alleles previously reported.^8,15 However, the peroxisome phenotype seems to be less dramatic in leaf mesophyll cells. For instance, in addition to the decreased number of total peroxisomes, peroxisomes in leaf cells are only slightly elongated or exhibit a beads-on-a-string phenotype (Fig. 1C and D). Previously, we reported the phenotypes of three strong drp3A alleles, all of which contain a large number of peroxules, long and thin membrane extensions from the peroxisome,⁸ yet such peroxisomal structures are not observed in pdd2. On the other hand, pdd2 has a more severe growth phenotype than most drp3A alleles, as it is slow in growth and has pale green leaves (Fig. 1E). Genetic analysis showed that pdd2 segregates as a single recessive mutation (data not shown).Open in a separate windowFigure 1Phenotypic analyses of pdd2 and identification of the PDD2 gene. (A–D) Confocal micrographs of root and mesophyll cells in 3-week-old wild type and pdd2 mutant plants. Green signals show peroxisomes; red signals show chloroplasts. Scale bars = 20 µm. (E) Growth phenotype of 3-week-old mutants. (F) Map-based cloning of the PDD2 gene. Genetic distance from PDD2 is shown under each molecular marker. Positions for mutations in previously analyzed drp3A alleles and pdd2 are indicated in the gene schematic. drp3A-1 and drp3A-2 are T-DNA insertion mutants, whereas pdd1 is an EMS mutant containing a premature stop codon in exon 6. (G) A schematic of the DRP3A (PDD2) protein with functional domains indicated. The pdd2 allele encodes a truncated protein lacking part of the GED domain.The unique combination of peroxisomal and growth phenotypes of pdd2 prompted us to use map-based cloning to identify the PDD2 gene, with the hope to discover novel proteins in the peroxisome division machinery. A population of approximately 6,000 F₂ plants (pdd2 × Ler) was generated. After screening 755 F₂ mutants, the pdd2 mutation was mapped to the region between markers T10C21 and F4B14 on the long arm of chromosome 4 (Fig. 1F). Since this region contains DRP3A, we sequenced the entire DRP3A gene in pdd2 and identified a G→A transition at the junction of the 18^th exon and intron (Fig. 1F). Further analysis revealed that the point mutation at this junction caused mis-splicing of intron 18, introducing a stop codon in the GTPase effector domain GED near the C terminus (Fig. 1G).DRPs share with the classic dynamins an N-terminal GTPase domain, a middle domain (MD), and a regulatory motif named the GTPase effector domain (GED) (Fig. 1G). To date, a total of 26 drp3A mutant alleles carrying missense or nonsense mutations along the length of the DRP3A gene have been isolated.^8,15 The combined peroxisomal and growth phenotype of pdd2 and the nature of the mutation in this allele are unique among all the drp3A alleles, indicating that the partial GED domain retained in pdd2 may have created some novel function for this protein. Further analysis of the truncated protein may be necessary to test this prediction.